Most cancer deaths are not caused by the primary tumor. Instead, death results from metastases, i.e., multiple widespread tumor colonies established by malignant cells that detach themselves from the site of the original tumor and travel through the body, often to distant sites. If a primary tumor is detected early enough, surgery, radiation, chemotherapy, or some combination of those treatments can often eliminate it. Unfortunately, the metastatic colonies are harder to detect and eliminate and it is often impossible to treat all of them successfully. Therefore, from a clinical point of view, metastasis can be considered the conclusive event in the natural progression of cancer. Moreover, the ability to metastasize is the property that uniquely characterizes a malignant tumor. Cancer metastasis comprises the following complex series of sequential events:    1. Extension from the primary locus into surrounding tissues;    2. Penetration into body cavities and vessels;    3. Release of tumor cells for transport through the circulatory system to distant sites;    4. Re-invasion of tissue at the site of arrest; and    5. Adaptations to the new environment so as to promote tumor cell survival, vascularization, and tumor growth.
Based on the complexity of cancer and cancer metastasis, and the frustration in treating cancer patients over the years, many attempts have been made to develop diagnostic tests to guide treatment and monitor the effects of such treatment on metastasis or relapse. Such tests presumably could also be used for cancer screening, replacing relatively crude tests such as mammography for breast tumors or digital rectal exams for prostate cancers. Towards that goal, a number of tests have been developed over the last 20 years and their benefits evaluated. One of the first attempts was the formulation of an immunoassay for carcinoembryonic antigen [CEA]. This antigen appears on fetal cells and reappears on tumor cells in certain cancers. Extensive efforts have been made to evaluate the usefulness of testing for CEA as well as many other “tumor” antigens, such as PSA, CA 15.3, CA125, PSMA, CA27.29. These efforts have proven to be somewhat futile as the appearance of such antigens in a test sample have not been generally predictive and are often detected when there is little hope for the patient. In the last few years, however, one test has proven to be useful in the early detection of cancer, viz., PSA for prostate cancers. When used with follow-up physical examination and biopsy, the PSA test has played a remarkable role in detecting prostate cancer early, at the time when it is best treated.
Despite the success of PSA testing, the test leaves much to be desired. For example, high levels of PSA do not always correlate with cancer nor do they appear to be an indication of the metastatic potential of the tumor. This may be due in part to the fact that PSA is a component of normal prostate tissue as well as other unknown factors. Moreover, it is becoming clear that a large percentage of prostate cancer patients will continue to have localized disease which is not life threatening. Based on the desire to obtain better concordance between those patients with cancers that will metastasize and those that will not, attempts have been made to determine whether prostate cells are in the circulation. When added to high PSA levels and biopsy data, the existence of circulating tumor cells might give indications as to how vigorously the patient should be treated.
The recommended approach for determining the presence of circulating prostate tumor cells has been to test for the expression of messenger RNA of PSA in blood. This is being done through the laborious procedure of isolating all of the mRNA from a blood sample and performing RT-PCR. No good correlation exists between the presence of such cells in blood and the ability to predict which patients are in need of vigorous treatment (L G Gomella, J of Urology, 158:326-337(1997)). It is noteworthy that PCR is difficult, if not impossible in many situations, to perform quantitatively, i.e., to determine the number of tumor cells per unit volume of biological sample. Additionally, false positives are often observed using this technique. An added drawback is the finite and practical limit to the sensitivity of this technique based on the sample size examined. Typically, the test is performed on 105 to 106 cells purified away from interfering red blood cells. With 5-10×106 leukocytes in normal blood, this corresponds to a practical lower limit of sensitivity of one tumor cell/0.1 ml of blood. Hence, there needs to be about 10 tumor cells in one ml of blood before signal is detectable. As a further potential complication, tumor cells are often genetically unstable. Accordingly, cancer cells having genetic rearrangements and sequence changes may be missed in a PCR assay as the requisite sequence complementarity between PCR primers and target sequences can be lost.
In summary, a useful diagnostic test needs to be highly sensitive and reliably quantitative. Such a test should be capable of detecting the presence of a single tumor cell in one ml of blood, thus corresponding on average, to 3000-4000 total cells in circulation. In inoculum studies for establishing tumors in animals that number of cells can indeed lead to the establishment of a tumor. Further, if 3000-4000 circulating cells represent 0.01% of the total cells in a tumor, then it would contain about 4×107 total cells. A tumor containing that number of cells would not be visible by any technique currently in existence. Hence, if tumor cells were shed in the early stages of cancer, a test with the sensitivity mentioned above would detect the cancer. If tumor cells were shed in some functional relationship with tumor size, then a quantitative test would be beneficial to assessing tumor burden. Heretofore, there has been no information reported regarding the existence of circulating tumor cells in very early cancers. Further, there are very considerable doubts in the medical literature regarding the existence of such cells and the potential of such information. The general view is that tumors are initially well confined and hence there will be few if any circulating cells in early stages of disease, and that early detection of cancer cells in circulation, even if feasible, would be unlikely to yield any useful information.
Based on the above, it is apparent that a method for identifying those cells in circulation with metastatic potential prior to establishment of a secondary tumor is highly desirable, particularly during the early stages of cancer. To appreciate the advantage such a test would have over conventional immunoassays, consider that a highly sensitive immunoassay has a lower limit of functional sensitivity of 10−17 moles. If one tumor cell can be captured from one ml of blood and analyzed, the number of moles of surface receptor, assuming 100,000 receptors per cell would be 10-19 moles. Since about 300 molecules can be detected on a cell, such an assay would have a functional sensitivity on the order of 10−22 moles, which is quite remarkable. To achieve that level of sensitivity in the isolation of such rare cells and to isolate them in a fashion which does not compromise or interfere with their characterization is a formidable task.
The introduction of flow cytometry to discriminate between cell populations has significantly improved the ability to accurately identify and enumerate components of cell populations that cannot be distinguished by morphological features. A further improvement of the sensitivity of flow cytometric examination of heterogeneous cell mixtures has been obtained by multidimensional analysis of the data. Cell populations are identified by the simultaneous assessment of light scattering and fluorescence parameters. Light scattering parameters measure cell size and cell granularity. Fluorescence parameters can be used to assess cell surface antigens, intracellular antigens, DNA, RNA, and protein content. By simultaneous analysis of light scatter and fluorescence parameters of individual cells passing through the laser beam, a multidimensional space is created in which the cells with dissimilar properties emerge in different locations. Conditions needed to detect infrequent/rare cells by flow cytometry are:
1. Sufficient sample volume for analysis;
2. Analysis by flow cytometry in a reasonable amount of time;
3. Selection of parameters such that the cell population of interest is located in a unique position;
4. Frequency of the target cells should be higher then 1 in 105 cells.
The current sample preparation procedures in which blood samples are incubated with fluorescently-labeled antibodies followed by addition of an erythrocyte lysing agent dilutes the sample ten-fold and is thus not suitable for detection of rare cells. In research laboratories, density separations or erythrocyte lysing procedures achieve reduction of the sample volume and an increase in cell concentration. These procedures lead to variable cell losses and are difficult to standardize between laboratories. Moreover, no significant enrichment of the target cells is obtained.
One method for isolating circulating tumor cells for analysis and enumeration is described in U.S. Pat. No. 6,365,362. The method described therein uses a magnetic particle labeled with antibodies to markers commonly found on circulating tumor cells that can be magnetically selected from a patient blood sample. Assays based on this method have shown not only that breast cancer tumor cells can be found in the blood of a patient with tumors at the lower limit of detection by mammography, but that the number of circulating tumor cells can be correlated to conventional therapies. For example, the number of circulating tumor cells decreases with chemotherapeutic treatments or surgery. Other tests using this method have shown that the number of circulating tumor cells is proportional to the tumor mass in several patients with colon cancer. Still, other tests showed that as a cancer patient comes out of remission, the number of circulating tumor cells increases. These remarkable results were found in a variety of cancers, including cancers of the breast, prostate, and colon.
As exciting as these results are, they must be tempered with the proper amount of scientific restraint. While the detection of circulating tumor cells in one's blood is frightening to the patient, a negative test result has not yet been proved to be an indication that a patient is free of cancer. Even worse would be a false-negative result for circulating blood cells. Reagent failure, instrument failure and operator errors can all lead to erroneous negative results. As the cancer cells in blood are rare, (often less than 1 cell/ml of blood) the blood volumes needed to perform the test are restrictively large. The requirement for such a large volume of blood prohibits the use of additional blood samples for traditional external control purposes. As discussed by Terstappen in “Detection of infrequent cells in blood and bone marrow by flowcytometry”, Hematopoietic Stem Cell Therapy, ed. A. Ho, Marcel Dekker Inc. pp. 137-152, (2000) to test non-specific (negative) binding in non-rare, traditional cell detection assays, the number of cells counted is generally less than 1% of the starting cell population. In the actual test, the specific reagents detect a cell population generally larger than 1%, thus confirming that the reagents are actually working. This non-specific binding (NSB) would result in a cell count of 105 cells, if one started with 107 cells and a NSB of 1%. However, in cancer cell detection, the specific binding of 0 cells may be detected in a cancer-free patient and must be discriminated from the presence of 1-100 circulating tumor cells in a patient who is undergoing relapse. With 0 cells detected, one has no way of knowing whether the reagents and/or process are working. An internal/indwelling control for assessing each of the components used in the test is thus desirable.
In order to have the required certainty that a test result is valid, controls at a number of essential points in the process are necessary. The first essential point that needs control is the magnetic labeling step. With so few tumor cells in the test sample, it is vital that these cells be targeted by the antibody-bearing magnetic particles. Another point is the magnetic selection of the magnetically labeled targets, which includes aspirating the excess liquid and non-selected cells, and the further washing of the magnetic particle/cell complexes. Still another point in the procedure is the step of labeling with antigen specific fluorescent dyes, some of which target antigen present on the cellular surface, but some of which require the permeabilization of the cellular and/or nuclear membrane. Yet another point is the enumeration of the actual target cells. As described in U.S. Pat. No. 6,365,362, enumeration is performed by flow cytometry, but use of the system described in U.S. Pat. No. 6,623,983 or the system described in U.S. Pat. No. 5,985,153 may also be employed if desired. These patents are incorporated by references herein.
One example of an experimental control is the use of ‘isotopic dilution’ to determine yield in chemical reactions or purifications. In this procedure, a pure sample of the molecule or compound of interest is labeled with a radioactive isotope of one of the atoms in the molecule. A known amount of the isotopically labeled compound is added to starting material and the chemical reaction or isolation procedure is run. At the end of the process, the percentage of isotopically labeled compound is calculated. The comparison between the original starting materials and the final product allows a calculation of the yield or percentage recovery of the starting material. This type of control also allows for sophisticated analysis of which steps in a process result in the loss of product or low yields. Use of a genuine ‘isotopic dilution’ protocol is not possible in the isolation of biological materials, such as cells, especially tumor cells. However, the use of cells which are labeled with a characteristic marker to distinguish them from the target cells, and which behave in a manner which could be proven to be very similar to the target cells would be useful to impart some information about percentage recovery to the researcher.
The traditional controls for immunophenotyping of cells are isotype controls. In an isotype control, the test is run using a monoclonal antibody of the same isotype, same species, but directed against an irrelevant antigen. In the circulating tumor cell assay mentioned above, the monoclonal antibody on the magnetic particle is directed against the epithelial cell adhesion molecule (EpCAM). The clone used in the examples in this specification is a mouse antibody IgG1κ. The traditional isotype control for this particle should be a magnetic particle prepared identically, only now the particle is labeled with a mouse antibody IgG1κ, directed against an antigen that does not appear in humans, such as keyhole limpet hemocyanin (KLH). Cells selected after magnetic separation with this isotype antibody on the magnetic particle are non-specifically bound, and the number of non-specifically selected cells can be determined. The IgG1κ monoclonal antibody directed against the leukocyte antigen CD45 is labeled with fluorescein isothiocyanate (FITC). The traditional isotype control is a FITC-labeled monoclonal IgG1κ antibody directed against an antigen which is not expressed in humans, such as KLH. Cells selected after magnetic separation and stained with this FITC-labeled isotype antibody determine the background staining in the FITC channel. The monoclonal antibody directed against the cytokeratins 4, 5, 6, 8, 10, 13, and 18 is labeled with phycoerythin (PE). This antibody is a murine monoclonal antibody, IgG1κ. The traditional isotype control is a PE-labeled monoclonal IgG1κ antibody directed against an antigen that does not appear in humans, such as KLH. Cells selected after magnetic separation and stained with this PE-labeled isotype antibody determine the background staining in the PE channel. Thus, all the antibodies in the system would be identical to those in the patient sample, except for the specificity. Cell selection with these reagents would be run side-by-side with a patient sample, using an identical aliquot of patient blood. If multi-parameter flow cytometry were used for the final analysis, the results would show a population of cells and the gates for the detection of tumor cells [FITC−, PE+] can be selected.
To discriminate between the non-specifically staining cells and the non-specifically selected cells, an additional blood sample, free of tumor cells, would be run using the isotype control magnetic particle, the CD45-FITC and the isotype control PE. If multi-parameter flow cytometry were used, the FITC[+] cells would be the non-specifically selected leukocytes. The FITC[+], PE[+] cells would be the non-specifically selected and the non-specifically staining cells. Cells that are FITC[−], PE[+] would be non-specifically selected cells that were binding non-specifically to PE, but not to the FITC MAb, as the isotype of both antibodies is the same. This non-specific binding is due to the fluorochrome, and not the antibody, or changes caused by the conjugation. Roughly, the same number of leukocytes would also appear in the patient sample with specific reagents, which also would have been non-specifically selected, yet specifically stained. Differential analysis effectively removes these leukocytes from the test results, offering further assurance that any “tumor cells” detected in the test are actually circulating epithelial cells and not non-specifically bound blood cells.
A more accurate control would be to use the EpCAM FF, the CD45-FITC, and an isotype PE MAb. In a patient sample, the majority of the selected cells are non-specifically selected. These cells are recognized by the CD45-FITC MAb and can thus be enumerated and they represent the true non-specific selection by the EpCAM FF. The actual tumor cells will not be stained with CD45-FITC, nor with the isotype control PE antibody. However, as the frequency is extremely low, one cannot determine whether there are actually tumor cells in the patient sample.
Although the traditional isotype controls described above represent the types of controls appropriate to cell selection, they are not truly functional controls. First, the level of background varies considerably, depending on which antibody is chosen for an isotype control. Thus, the choice of isotype controls could be made to influence a higher or lower background threshold. Second, this type of control does not account for the reagents used in the actual patient test. For example, the anti-cytokeratin antibody in the patient test may have been inadvertently omitted from the test mixture. This mistake would be undetected by the isotype control. Finally, and most importantly, this type of isotype control can be used for small blood samples, which require 50-150 μl of blood. However, in rare cell isolation, a full tube, and optimally 5-30 ml of whole blood is required for the detection of tumor cells. Cell numbers as low as 1 cell/ml of blood have been detected, thus the larger the sample, the less likely the test will miss a patient with a low incidence of circulating tumor cells. These cancer patients are already subjected to a variety of medical tests, so the draining of an extra 20-30 ml of blood for an isotype control of limited value is not acceptable. Use of a small blood sample for an isotype control is also of limited value. Dividing the blood sample into an isotype control and the actual test sample decreases sensitivity, and those patients with a low level of circulating tumor cells would be missed. As documented by Stelzer, et al, Cytometry 30:214-230 (1997) and Keeney, et al, Cytometry 34:280-283 (1998), consensus is building towards elimination of a patient sample for use as any type of control, including an isotype control.
Immunicon's U.S. Pat. No. 5,985,153 describes an internal control, which is substantially different from an external, isotype control. In Examples 6 and 7 of the '153 patent, beads with a magnetic “load” or antigen “loads” similar to those found on cancer cells are added to the blood sample. The percentage or number of beads detected by the test is used to determine the efficiency of the test. The use of beads as a control is well known in the art and has a clear advantage as there is no chance of mistaking a bead for a cell during the analysis of the test. Beads also store well and can be reproducibly manufactured, and have the added benefit that they can be used to accurately determine the volume of a sample. As described by Stewart et al, Cytometry 2(4): 238-243 (1982), the use of a known quantity of fluorescent beads overcomes the common problem of determining the sample volume actually analyzed by a flow cytometer. However, the use of beads as a control has limitations. Beads can be extremely sturdy, and as such unaffected by numerous conditions that would destroy a cell, thus limiting their usefulness as a control against operator error. Sensitivity of the bead to conditions such as temperature, pH, and isotonic strength should be similar to that of an actual cell. The engineering and manufacturing considerations for forming such a bead with the appropriate antigens, antigen density, and dyes to provide a control for the cell selection would be difficult. Not only must the bead have the appropriate antigens, they must be accessible under conditions similar to that of an actual cell. For example, steric factors and binding constants must be taken into consideration. Finally, beads are solid objects, not affected by the permeabilization reactions, limiting their usefulness as a control at that crucial step. Therefore, even if one could perfectly duplicate a cell surface, beads could still not serve as a true internal, positive control for a cell selection test.
Another approach to providing a control would be to use actual cells as controls. Indeed, a standard quality control procedure for cell surface phenotyping is to obtain specimens from normal donors to be prepared and analyzed concomitantly with the patient's sample. Ideally, the normal specimen is of the same type, and obtained at the same time, as the patient sample, although this is generally not possible for specimens other than peripheral blood. Even with peripheral blood, the use of fresh blood can be costly, time-consuming, and not always available, causing many labs to turn to stored cell products as the source of their controls. The use of prepared, commercially obtained, preserved cells as controls for various medical tests are well known in the art. Control cells embedded in gelatin, paraffin, or agar are described in U.S. Pat. No. 5,610,022 (Battifora) and U.S. Pat. No. 5,187,099 (Healy, et al.). The use of preserved cells for reference controls for cell counters are described in U.S. Pat. No. 5,981,282 (Ryan); U.S. Pat. No. 5,432,089 (Ryan); U.S. Pat. No. 5,342,754 (Maples, et al); and U.S. Pat. No. 5,763,204 (Maples, et al.). Preservation by lyophilization is also used, as described in U.S. Pat. No. 5,059,518 (Kortright, et al.); U.S. Pat. No. 5,968,831 (Shukla, et al.); and European patent 469 766B1 (Davis). The creation of a standard solutions used for cells counters, flow cytometers and other instruments are described in U.S. Pat. No. 5,529,933 (Young, et al.); U.S. Pat. No. 5,888,823 (Matsumoto, et al.); and Japanese accepted specification 01259261. A review of the catalogs of the major suppliers of reagents for hematology analyzers, flow cytometers, and other cell analysis platforms reveals a large number of cell-based controls for these instruments. Some examples include Streck Laboratories Cell-Chex® and Chem-Chex reagents, R&D Systems R&D Retic reagents, BeckmanCoulter Cyto-Trol® control cells, and BioErgonomics FluoroTrol® line of stabilized leukocytes.
Although the control cells described in the above-mentioned patents and the various commercially available reagent lines offer many forms of stabilized cells for cell procedures, the methods and reagents described would only be able to provide external controls for cell selection and analysis procedure. None would provide a suitable internal control for the selection and enumeration of rare cells, such as circulating tumor cells. In addition, the stability of the cells is limited to 14-30 days. In those cases where there is longer stability, the cells have been lyophilized, which increases shelf life, but may decrease reproducibility, due to inadequate reconstitution.
If one were to use cells as an external or internal, positive control for rare cell selection, many problems are presented. For example, how are the cells obtained? How can the control cells be differentiated from the target cells? How can one prove that the control cells behave similar to the target cells? How can the cells be used to control the experiment? Is the recognition ability similar across all levels of target cell detection? Use, reproducibility, and efficacy are essential concerns regarding the use of cells as an external or internal positive control for cell selection, all of which are addressed in the present invention.